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GlomusafricanumandG.iranicum,twonewspeciesofarbuscularmycorrhizalfungi(Glomeromycota)

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Glomus africanum and G. iranicum, two new speciesof arbuscular mycorrhizal fungi (Glomeromycota)

Janusz Błaszkowski1

Department of Plant Protection, West PomeranianUniversity of Technology, Szczecin, Słowackiego 17,PL-71434 Szczecin, Poland

Gabor M. KovacsDepartment of Plant Anatomy, Institute of Biology,Eotvos Lorand University, Pazmany Peter setany 1/C,1117 Budapest, Hungary

Tımea K. BalazsInstitute of Ecology and Botany, Hungarian Academy ofSciences, Alkotmany street 2–4, 2163 Vacratot, Hungary

Elz_bieta OrłowskaInstitute of Molecular Biology, University of Aarhus,Gustav Wieds Vej 10 C, 8000 Aarhus C Denmark

Mehdi SadraviDepartment of Plant Protection, Faculty of Agriculture,Yasouj University, Daneshju Avenue, P.O. Box 353,75918–74831 Yasouj, Iran

Tesfaye WubetFrancois Buscot

UFZ, Helmholtz Centre for Environmental Research,Theodor-Lieser-Straße 4, 06120 Halle-Saale, Germany

Abstract: Two new arbuscular mycorrhizal fungalspecies (Glomeromycota) of genus Glomus, G. africa-num and G. iranicum, are described and illustrated.Both species formed spores in loose clusters andsingly in soil and G. iranicum sometimes inside roots.G. africanum spores are pale yellow to brownishyellow, globose to subglobose, (60–)87(–125) mmdiam, sometimes ovoid to irregular, 80–110 3 90–140 mm. The spore wall consists of a semipermanent,hyaline, outer layer and a laminate, smooth, paleyellow to brownish yellow, inner layer, which always ismarkedly thinner than the outer layer. G. iranicumspores are hyaline to pastel yellow, globose tosubglobose, (13–)40(–56) mm diam, rarely egg-shaped, prolate to irregular, 39–54 3 48–65 mm.The spore wall consists of three smooth layers: onemucilaginous, short-lived, hyaline, outermost; onepermanent, semirigid, hyaline, middle; and onelaminate, hyaline to pastel yellow, innermost. Onlythe outermost spore wall layer of G. iranicum stains

red in Melzer’s reagent. In the field G. africanum wasassociated with roots of five plant species and anunrecognized shrub colonizing maritime sand dunesof two countries in Europe and two in Africa, and G.iranicum was associated with Triticum aestivumcultivated in southwestern Iran. In one-species cul-tures with Plantago lanceolata as the host plant G.africanum and G. iranicum formed arbuscular mycor-rhizae. Phylogenetic analyses of partial SSU sequencesof nrDNA placed the two new species in Glomusgroup A. Both species were distinctly separated fromsequences of described Glomus species.

Key words: arbuscular fungi, Glomeromycota,molecular phylogeny, mycorrhizae, new species

INTRODUCTION

Arbuscular mycorrhizal fungi (AMF) of phylumGlomeromycota are the most common soil fungi inthe world coexisting symbiotically with ca. 70–90% ofland plants (Wang and Qiu 2006, Smith and Read2008, Brundrett 2009). Maritime sand dunes favorAMF development (Koske 1987, Dalpe 1989, Tadychand Błaszkowski 2000) because of low nutrient andorganic matter content (Nicolson and Johnston 1979,Koske 1988), as well as the absence of numerousantagonistic microorganisms, especially parasites ofAMF (Koske et al. 2004).

Of the ca. 220 described species of Glomeromycota,at least 35 originally were isolated from maritimedunes and many others have been associated withroots of dune plants (Sridhar and Beena 2001, www.agro.ar.szczecin.pl/,jblaszkowski/). Members of Glo-meromycota also commonly co-occur with cultivatedplants, including Triticum aestivum L. that usually hasharbored abundant and diverse spore populations ofthese fungi (Hetrick and Bloom 1983, Dodd andJeffries 1989, Błaszkowski 1993).

AMF sequences amplified from root sampleshowever suggest that the number of existing speciesof AMF is much higher than that formally describedand that most undescribed species belong to genusGlomus (Helgason et al. 2002, Fitter 2005, Hijri et al.2006, Kovacs et al. 2007, Opik et al. 2009), especiallyin Glomus group A sensu Schwarzot et al. (2001).Possible causes for the omission of these undescribedspecies from the scientific record might be due to (i)a lack of or rare sampling of AMF in many terrestrialregions of Earth, (ii) the few specialized and

Submitted 3 Dec 2009; accepted for publication 14 Apr 2010.1 Corresponding author. Department of Plant Protection, WestPomeranian University of Technology, Szczecin, Słowackiego 17, PL71434 Szczecin, Poland. E-mail: janusz.blaszkowski@zut.edu.pl

Mycologia, 102(6), 2010, pp. 1450–1462. DOI: 10.3852/09-302# 2010 by The Mycological Society of America, Lawrence, KS 66044-8897

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experienced mycologists that study taxonomy ofGlomeromycota, and (iii) seasonal, rare or lack ofsporulation by many AMF in the field (Gemma et al.1989, Sturmer and Bellei 1994, Stutz and Morton1996). Often the diversity of AMF spores obtainedfrom environmental samples can be increased withsuccessive (Stutz and Morton 1996) or long-term(Oehl et al. 2004) pot trap cultures.

Examination of long-term trap cultures with rhizo-sphere soils and roots of plant species collected frommaritime sand dunes of Africa and Europe and fromT. aestivum cultivated in southwestern Iran revealedspores of two undescribed species of Glomeromycotaforming glomoid spores. Phylogenetic analyses ofsequences of rDNA placed the fungi in Glomus groupA sensu Schwarzot et al. (2001) and confirmed theiruniqueness relative to other known Glomus species.The fungi are described here as G. africanum sp. nov.and G. iranicum sp. nov.

MATERIALS AND METHODS

Establishment and growth of trap and single-species cultures,extraction of spores and staining of mycorrhizae.—Sporesexamined in this study were derived from both pot trap andsingle-species cultures. Trap cultures were established toobtain a large number of living spores and to initiatesporulation of species that were present but were notdetected in field collections (Stutz and Morton 1996). Themethod used to establish trap cultures, growing conditionsand the methods of spore extraction and staining ofmycorrhizae were those described by Błaszkowski et al.(2006).

Single-species cultures also were established and grown asdescribed by Błaszkowski et al. (2006), with three excep-tions. First, cultures of both species were successfullyestablished from small clusters of spores. The clustersconsisted of 2–3 (G. africanum) or 10 (G. irranicum) sporesattached by a common mycelium. To prevent contamina-tion by fragments of hyphae of other AMF the clusters wererinsed several times with water; each time the water wasremoved with a pipette. Second, instead of marine sand, thegrowing medium was an autoclaved commercially availablecoarse-grained sand (grains 1.0–10.0 mm diam, 80.50%;grains 0.1–1.0 mm diam, 17.28%; grains , 0.1 mm diam,2.22%) mixed (5 : 1, v/v) with clinopthilolite (Zeocem,Bystre, Slovakia) of grains 2.5–5 mm. Clinopthilolite is acrystaline hydrated alumosilicate of alkali metals andalkaline earth metals having a high ion exchange andwater-holding capacity. The pH of the sand-clinopthilolitemixture was 7.3. Third, the cultures were kept in transpar-ent plastic bags, 15 cm wide and 22 cm high, as suggested byWalker and Vestberg (1994), instead of open pot cultures(Gilmore 1968). To prevent contamination of cultures withother AMF but still allow gas exchange an opening of about1 cm2 was left in the upper part of each bag while the edgeswere sealed with plastic clips. The cultures were wateredwith tap water once a week and harvested after 5 mo to

extract spores. Root fragments located ca. 1–5 cm below theupper level of the growing medium were cut off with ascalpel to reveal mycorrhizal structures. Plantago lanceolataL. was used as a host plant in both trap and single-speciescultures.

Microscopy survey.—Morphological properties of spores andwall structure were determined based on examination of atleast 100 spores mounted in water, lactic acid, polyvinylalcohol/lactic acid/glycerol (PVLG, Omar et al. 1979) anda mixture of PVLG and Melzer’s reagent (1 : 1, v/v). Sporesat all developmental stages were crushed to varying degreesby applying pressure to the cover slip and then stored at 65 Cfor 24 h to clear contents from oil droplets. They wereexamined under an Olympus BX 50 compound microscopeequipped with Nomarski differential interference contrastoptics. Microphotographs were recorded on a Sony 3CDDcolor video camera coupled to the microscope.

Terminology of spore structure is that suggested bySturmer and Morton (1997) and Walker (1983). Sporecolor was examined under a dissecting microscope on freshspecimens immersed in water. Color names are fromKornerup and Wanscher (1983). Nomenclature of fungiand plants is that of Walker and Trappe (1993) and Mirek etal. (1995), respectively. The authors of the fungal names arethose presented at the Index Fungorum Website http://www.indexfungorum.org/AuthorsOfFungalNames.htm.Voucher specimens were mounted in PVLG and a mixtureof PVLG and Melzer’s reagent (1 : 1, v/v) on slides anddeposited in the Department of Plant Protection (DPP),West Pomeranian University of Technology, Szczecin,Poland, and in the herbarium at Oregon State University(OSC) in Corvallis, Oregon. Color microphotographs ofspores of the new species can be viewed at the URL http://www.agro.ar.szczecin.pl/,jblaszkowski/.

DNA extraction, polymerase chain reaction andDNA sequencing.—Several spores or small spore clusterswere used to obtain target DNA as described by Błaszkowskiet al. (2009). We used a nested PCR to amplify a segment ofSSU of the nrDNA. The GlomerWT0 and Glomer1536primers were used in the first PCR as described by Wubet etal. (2006). In the second step we used the AML1 and AML2primers as described by Lee et al. (2008). A high fidelityenzyme mix (MBI Fermentas, Vilnius, Lithuania) was usedfor PCR. With these primer combinations we obtainedsequences longer than 700 nucleotides, which are suitablefor reliable phylogenetic analyses of Glomus groups.Because the amplified region overlaps the segment ampli-fied with AM1 (Helgason et al. 1998) and NS31 (Simon etal. 1992), primers used most widely in AMF diversity studies,we could compare the sequences of the new taxa withnumerous environmental AMF sequences available inpublic databases. The appropriate size amplicons werecleaned and either cloned into a pGEMT-easy vector(Promega, Madison, Wisconsin) and transformed intocompetent JM109 Escherichia coli (Promega, Madison,Wisconsin) or cloned with the TOPO TA CloningH Kit(Invitrogen) and transformed into TOP10 chemicallycompetent E. coli strains (Invitrogen) following manufac-turers’ instructions. Ten positive clones from both species

BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1451

were sequenced in both directions with universal primersand an ABI PRISM 3.1 BigDye Terminator 3.1 CycleSequencing Kit (Applied Biosystems, Foster City, Califor-nia). Electrophoreses were carried out on an ABI PRISM3100 or 3730XL Genetic Analyzer (Applied Biosystems,Foster City, California). The electrophoregrams wereprocessed with Pregap4 1.4b1 and Gap 4.8b1 programs ofthe Staden Program Package (Staden et al. 2000). Nonre-dundant sequences of clones were deposited in GenBank(HM153415-HM153424).

Phylogenetic analyses.—After pilot analyses of the sequencestogether with identified species of the phylum Glomeromy-cota the final analyses were carried out with a dataset ofknown Glomus group A sequences and unidentified AMFsequences from in planta studies including the most similarsequences to our clones obtained from BLAST queries. Weused Glomus lamellosum as outgroup. Only the two mostdistant sequences of both new taxa were included in theanalyses. The sequences were aligned with Multalin (Corpet1988, http://prodes.toulouse.inra.fr/multalin/multalin.html) and manually edited with ProSeq 2.9 (Filatov 2002).The best fit nucleotide substitution model was selected withthe program jModelTest (Posada 2008) considering theselection of Akaike information criterion (AIC). The modeland the parameters were used to calculate distances forneighbor-joining analyses with PAUP*4.0b10 software(Swofford 2003). Support of branches was tested bybootstrap analysis with 1000 replicates. A maximumlikelihood (ML) phylogenetic analysis was carried out withthe online version of PHYML 3.0 (Guindon and Gascuel2003). The GTR nucleotide substitution model was usedwith ML estimation of base frequencies. The proportion ofinvariable sites was estimated and optimized. Six substitu-tion rate categories were set, and the gamma distributionparameter was estimated and optimized. A bootstrapanalysis with 1000 replicates also was used here to testsupport of branches. The same substitution model was usedin Bayesian analyses performed with MrBayes 3.1 (Huelsen-beck and Ronquist 2001, Ronquist and Huelsenbeck 2003)with the Computational Biology Service Unit, CornellUniversity (http://cbsuapps.tc.cornell.edu/index.aspx).The Markov chain was run 5 000 000 generations, samplingin every 100 steps, and with a burn-in at 7500 sampled trees.The alignment was deposited in TreeBase (http://purl.org/phylo/treebase/phylows/study/TB2:S10461). Phyloge-netic trees were viewed and edited by Tree Explorer of theMEGA 4.0 program (Tamura et al. 2007) and a text editor.

TAXONOMY

Glomus africanum Błaszk. & Kovacs sp. nov. FIGS. 1–13MycoBank MB518241

Sporocarpia ignota. Sporae singulatim vel gregatim insolo efformatae. Fascicula 470–620 3 600–1250 mm, e sporis2–6. Sporae pallide luteae vel spadiceae; globosae velsubglobosae; (60–)87(–125) mm diam; raro ovoideae,oblongae vel irregulares; 80–110 3 90–140 mm. Tunicasporae stratis duobus (strata 1 ad 2); stratum ‘‘1’’ caducum,glabrum, hyalinum, (1.5–)3.1(–8.6) mm crassum; stratum

‘‘2’’ laminatum, glabrum, pallide luteum vel spadiceum,(1.0–)1.7(–2.7) mm crassum. Hypha sporifera pallide luteavel spadicea; recta vel recurvta; cylindrica vel infundibuli-formis; (3.7–)5.7(–9.3) mm lata ad basim sporae; parietepallide luteo vel spadiceo, (2.9–)4.4(–5.4) mm crasso, stratis1 ad 2 in parietem sporae continuantibus. Porus hyphae(1.0–)2.1(–2.9) diam. Mycorrhizas vesiculo-arbuscularesformans.

Typus: Polonia: Sedinum (Szczecin), infra P.lanceolata, 10 Mar 2008, J. Błaszkowski, 3167 (Holo-typus, DPP).

Sporocarps unknown. Spores formed in loose clus-ters or singly in the soil (FIGS. 1, 2) develop blasticallyat the tip of sporogenous hyphae either branchedfrom a parent hypha continuous with a mycorrhizalextraradical hypha (spores in clusters) or directlydeveloped from mycorrhizal extraradical hyphae(single spores). Clusters 470–620 3 600–1250 mm with2–6 spores (FIG. 1). Spores pale yellow (4A3) tobrownish yellow (5C8); globose to subglobose,(60–)87(–125) mm diam, sometimes ovoid to irregular;80–110 3 90–140 mm; with one subtending hypha(FIGS. 1–3, 7 and 8). Spore wall composed of two layers(FIGS. 3–5, 7, 8). Layer 1, forming the surface,semipermanent, evanescent, hyaline, (1.5–)3.1(–8.6)mm thick, more or less deteriorated in mature spores,infrequently completely sloughed in older specimens;in young and freshly matured spores, the uppersurface of this layer usually is covered with irregularblister-like outgrowths, rarely is smooth (FIGS. 3–8).Layer 2-laminate, smooth, pale yellow (4A3) tobrownish yellow (5C8), (1.0–)1.7(–2.7) mm thick(FIGS. 3–5, 7, 8). Layers 1 and 2 do not stain inMelzer’s reagent. Subtending hypha pale yellow (4A3)to brownish yellow (5C8); straight or recurved, flaredto slightly funnel-shaped, sometimes slightly constrict-ed at spore base; (3.7–)5.7(–9.3) mm wide at the sporebase (FIGS. 1–3, 7, 8). Wall of subtending hypha paleyellow (4A3) to brownish yellow (5C8); (2.9–)4.4(–5.4)mm thick at spore base; continuous with spore walllayers 1 and 2; layers 1 and 2 usually extend far belowspore base in mature spores (FIGS. 2, 3, 7, 8). Pore(1.0–)2.1(–2.9) mm diam, open (FIG. 7) or occluded bya curved septum continuous with some innermostlaminae of spore wall layer 2 (FIG. 8). Germinationunknown.

Mycorrhizal associations. In the field G. aficanumwas associated with roots of Ammophila arenaria (L.)Link, Cineraria geifolia L., Senecio elegans L., Thino-pyrum distichum (Thunb.) A. Love, Trachyandradivaricata ( Jacq.) Kunth and an unrecognized shrub.

In one-species culture with P. lanceolata as the hostplant G. africanum formed mycorrhizae with arbus-cules, vesicles and intra- and extraradical hyphae(FIGS. 9–12). Arbuscules generally were dispersedwidely along the root fragments examined. They

1452 MYCOLOGIA

FIGS. 1–8. Glomus africanum. 1. Spores in loose cluster. 2. Single spore. 3–5. Spore wall layers (swl) 1–2; note the irregularblister-like outgrowths covering the upper surface of swl1 seen in a cross view. 6. Irregular outgrowths of the spore surface seenin a plan view. 7. Subtending hyphal wall layers (shwl) 1 and 2 continuous with spore wall layers (swl) 1 and 2; note the openlumen of the subtending hypha. 8. Subtending hyphal wall layers (shwl) 1 and 2 continuous with spore wall layers (swl) 1 and2; note the septum in the lumen of the subtending hypha. 1, 2. Spores in lactic acid. 4, 6–8. Spores crushed in PVLG. 3, 5.Spores in PVLG + Melzer’s reagent. 1–8, differential interference microscopy. Bars: 1, 2 5 20 mm; 3–8 5 10 mm.

BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1453

consisted of a short trunk grown from a parent hyphaand numerous branches with fine tips (FIG. 9).Vesicles were not numerous and usually highlyseparated. They were ellipsoidal to oblong, 10.5–34.5 3 25.5–113.8 mm, when observed in a plan view(FIG. 10). Intraradical hyphae grew along the rootaxis, were (1.3–)4.6(–9.8) mm wide, straight or slightlyrecurved, and occasionally formed H- or Y-shapedbranches and coils (FIGS. 9–12). The coils wereellipsoidal to oblong, 14.0–21.6 3 50.0–82.4.0 mm,when seen in a plan view (FIG. 11). Extraradicalhyphae were (2.5–)4.1(–5.5) mm wide and occurredinfrequently. In 0.1% trypan blue arbuscules stainedviolet white (17A2) to violet (17B6), vesicles pastelviolet (17A4) to deep violet (17D8), intraradicalhyphae pale violet (17A3) to violet (17B8), coilspastel violet (17A4) to deep violet (17D8), andextraradical hyphae pale violet (17A3) to deep violet(17D8; FIGS. 9–12).

Phylogenetic position. Phylogenetic analyses of par-tial SSU sequences of nrDNA placed G. africanumunambiguously in Glomus group A sensu Schwarzott et

al. (2001) within genus Glomus (FIG. 13). Thesequences of the species separated unambiguouslyfrom described Glomus species of this group. Sequenc-es of G. africanum showed high similarity to andgrouped together with in planta sequences fromJuniperus procera Hochst. ex Endlicher, Podocarpusfalcatus (Thunb.) R.Br. ex Mirb. and Prunus africanaHook.f. trees of the Afromontane region of Ethiopia(Wubet et al. 2006, 2009). This clade formed a sistergroup of species with subgroup ‘‘a’’ of Glomus group Asensu Schwarzott et al. (2001) with strong bootstrap(NJ 99%, ML 98%) and posterior probability (PP100%) support values. The high similarity of G.africanum to AMF sequences obtained from Africa(Wubet et al. 2006, 2009) is especially interestingbecause the species first was found on that continent(see below), although from a completely differenthabitat (maritime dune). Other environmental AMForiginating from different regions and continents alsoshowed high similarity to G. africanum when theBLAST query was restricted to the AM1-NS31 segmentof the SSU sequences (data not shown).

FIGS. 9–12. Mycorrhizae of Glomus africanum in roots of Plantago lanceolata stained in 0.1% trypan blue. 9. Arbuscule(arb) with trunk (t) developed from parent hypha (ph). 10. Vesicles (ves). 11. H-shaped branch (Hb). 12. Y-shaped branch(Yb). 9–12. Differential interference microscopy. Bars: 9, 11, 12 5 10 mm; 10 5 20 mm.

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FIG. 13. Neighbor-joining tree showing the phylogenetic positions of Glomus africanum and G. iranicum within Glomusgroup A inferred from 71 nrDNA SSU sequences with Glomus lamellosum as outgroup. The sequences obtained in this studyare shown in boldface. Geographic origin (in parentheses), GenBank accession numbers and hosts of in planta sequencesfrom earlier studies are provided. Values above branches have NJ bootstrap values (1000 replicates; before the slash) and MLbootstrap values (1000 replicates; after slash) as percentages, whereas values below branches are posterior probabilitiescalculated by Bayesian analysis as percentages. Bootstrap values below 75% and posterior probabilities below 90% are notshown. Bar 5 1 change/100 characters.

BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1455

Specimens examined. POLAND, Szczecin, under pot-cultured P. lanceolata, 10 May 2009, Błaszkowski, J., 3167(HOLOTYPE, DPP); Błaszkowski, J., 3168–3184 (ISTO-TYPES, DPP) and two slides at OSC.

Etymology. Latin, africanum, referring to the conti-nent from where the fungus was first found.

Distribution and habitat. With traditional methodsof finding AMF (not molecular) G. africanum hasbeen isolated from six trap cultures containingmixtures of rhizosphere soils/root fragments of fourrecognized plant species and an unrecognized shrubfrom two African countries (Egypt, South Africa) andfrom two trap cultures with soils and roots collectedunder A. arenaria growing in Bulgaria and Poland(Europe). All the plants colonized maritime sanddunes. No spores of AMF were isolated directly fromfield-collected samples. The South African plantspecies sampled were C. geifolia, S. elegans, Tr.divaricata, growing near Strand (34u069S, 18u499E),ca. 50 km southeast of Cape Town and Th. distichum,growing near Strand and in the Reserve Rooiels(34u189S, 18u499E). Strand samples were collected 31Jul–2 Oct 2005 and those from the Reserve Rooiels 2Aug 2005. The unrecognized shrub was sampled fromGiftung Island (27u109N, 33u569E), Egypt. The rhizo-sphere soil-root mixture of this plant was sampled 28Jul 2007. A. arenaria was sampled from dunes of theBlack Sea near Varna (43u139N, 27u559E), Bulgaria, on15 Sep 1998 and from dunes of the Baltic Sea adjacentto Swinoujscie (53u559N, 14u149E), northwestern Po-land, 10 Jul 2006.

Spores of G. africanum were not found in ca. 3000field-collected soils or in ca. 2500 pot trap culturesrepresenting other regions of Africa and Europe aswell as Asia and USA (Błaszkowski pers obs).

Notes. The most distinctive structures of G. africa-num are its two spore wall layers (FIGS. 3–8), of whichthe outer layer is hyaline, irregular and much thickerthan the structural laminate inner layer, which isexceptionally thin compared with the thickness of thelaminate structural spore wall layer of other knownGlomus spp. (www.agro.ar.szczecin.pl/,jblaszkowski/).

Of the species of Glomeromycota forming glomoid-colored spores with a two-layered spore wall inwhich the inner layer is laminate, G. africanum sporesmost resemble in color and size those of G.etunicatum W.N. Becker & Gerd. and G. versiforme(P. Karsten) S.M. Berch. However the darkest sporesof G. africanum are markedly darker than thedarkest G. versiforme spores (www.agro.ar.szczecin.pl/,jblaszkowski/, Błaszkowski et al. 2003). MoreoverG. versiforme spores may be (i) produced singly and incompact epigeous sporocarps (vs. singly and in looseclusters for G. africanum; FIGS. 1, 2) and (ii) slightlylarger, (80–)106(–150) mm diam when globose

(Daniels and Trappe 1979, Błaszkowski et al. 2003,www.invam.caf.wvu.edu/). Glomus etunicatum pro-duces only single hypogeous spores (Becker and Ger-demann 1977, www.agro.ar.szczecin.pl/,jblaszkowski/,www.invam.caf.wvu.edu/).

Both spore wall layers of G. africanum and G.versiforme are of the same type and do not stain inMelzer’s reagent. However spore wall layer 1 of G.africanum is much thicker and layer 2 much thinnerthan layer 1, (0.7–)1.0(–1.2) mm thick, and layer 2,(2.7–)4.1(–5.4) mm thick, of the spore wall of G.versiforme (www.agro.ar.szczecin.pl/,jblaszkowski/,Błaszkowski et al. 2003).

The semipermanent spore wall layer 1 of G.africanum is relatively long-lived and nonreactive inMelzer’s reagent (FIGS. 2–8), while that of G.etunicatum is short-lived, mucilaginous and stains inthis reagent (www.agro.ar.szczecin.pl/,jblaszkowski/,Sturmer and Morton 1997). In addition spore walllayer 1 of G. etunicatum is much thinner, 0.5–2.5 mmthick when intact, than spore wall layer 1 of G.africanum, and the upper range of thickness of thelaminate spore wall layer 2 of the latter species does notattain even the lower limit of the range of thickness ofthe laminate spore wall layer of the former fungus,4.5 mm thick (www.agro.ar.szczecin.pl/,jblaszkowski/).Finally, the subtending hypha of G. africanum sporesis less regular (flared to slightly funnel-shaped;FIGS. 1–3, 7, 8) than that of spores of both G.etunicatum and G. versiforme (cylindrical to flared;www.agro.ar.szczecin.pl/,jblaszkowski/, Błaszkowskiet al. 2003, www.invam.caf.wvu.edu/).

Molecular-phylogenetic analyses results (FIG. 13)indicate that G. africanum has no apparent relativesamong described Glomus spp. The phylogeneticposition of G. africanum within Glomeromycota isdifferent than that of G. etunicatum and G. versiforme;Glomus africanum has grouped among members ofGlomus group A, whereas G. etunicatum representsGlomus group B (Schwarzott et al. 2001) and Glomusversiforme is a close relative of Diversispora spurca(C.M. Pfeiff., C. Walker & Bloss) C. Walker &Schuessler, the type species of family DiversisporaceaeC. Walker & Schuessler (Walker and Schubler 2004,Redecker et al. 2007).

Of members of Glomus group A, juvenile G.constrictum Trappe spores are similar to mature,small-spored isolates of G. africanum in color andappearance. Moreover spore wall layer 2 of immatureG. constrictum spores is of thickness similar to that oflayer 2 of mature G. africanum spores (Błaszkowskipers obs). However at maturity G. constrictum sporesare much darker, brownish orange (6C8) to darkbrown (9F5) to black, than those of G. africanum, andspore wall layer 2 of the former species always is

1456 MYCOLOGIA

thicker, (7.5–)10.0(–12.0) mm, than layer 1, (0.8–)2.5(–8.5) mm thick, and much thicker than spore walllayer 2 of G. africanum (Trappe 1977, www.agro.ar.szczecin.pl/,jblaszkowski/). In addition G. constric-tum spores generally are much larger, (100–)160(–220–330) mm diam when globose, than those of G.africanum and the width of the subtending hypha ofspores of the former species far exceeds that of thelatter fungus, (11.3–)15.0(–17.5) mm wide at the sporebase in G. constrictum. Finally, while the subtendinghypha of G. constrictum spores typically is markedlyconstricted at the spore base (Trappe 1977; www.agro.ar.szczecin.pl/,jblaszkowski/), that of G. africanumspores is rarely and only slightly constricted at thebase.

Glomus iranicum Błaszk., Kovacs & Balazs, sp. nov.FIGS. 13–25

MycoBank MB518242Sporocarpia ignota. Sporae singulatim vel gregatim in

solo vel in radice efformatae. Fascicula globosa, oblonga velirregulares 70–280 3 90–480 mm. Sporae hyalinae velsubluteae; globosae vel subglobosae; (13–)40(–56) mmdiam; raro ovoideae, oblongae vel irregulares; 39–54 3

48–65 mm. Tunica sporae stratis tribus (strati 1–3); stratum‘‘1’’ caducum, glabrum, hyalinum, (0.4–)1.0(–1.5) mmcrassum, in solutione Melzeri rufum; stratum ‘‘2’’ semirigi-dum, glabrum, hyalinum, (0.8–)1.2(–1.5) mm crassum;stratum ‘‘3’’ laminatum, glabrum, hyalinum vel subluteum,(1.2–)2.0(–2.6) mm crassum. Hypha sporifera hyalina; rectavel recurvta; cylindrica vel infundibuliformis; (4.8–)6.9(–9.8) mm lata ad basim sporae; pariete hyalino vel subluteo,(1.8–)2.6(–3.5) mm crasso, stratis 1–3 in parietem sporaecontinuantibus. Porus hyphae (1.2–)2.5(–5.0) diam, aperto.Mycorrhizas arbusculares formans.

Typus: Polonia: Sedinum (Szczecin), infra P.lanceolata, 10 Mar 2008, J. Błaszkowski, 3185 (Holo-typus, DPP).

Sporocarps unknown. Spores formed in the soil inloose to compact clusters (FIGS. 14–17 and 22); 70–280 3 90–480 mm; rarely singly (FIG. 18), occasionallyinside roots; develop blastically at the tip ofsporogenous hyphae branched from a parent hyphacontinuous with a mycorrhizal extraradical hypha(FIGS. 14, 15), rarely intercalary. Spores hyaline topastel yellow (3A4), globose to subglobose, (13–)40(–56) mm diam, rarely egg-shaped, prolate toirregular; 39–54 3 48–65 mm; with one subtendinghypha (FIGS. 14–18 and 20–22). Spore wall composedof three layers (1–3, FIGS. 16–22). Layer 1, formingthe spore surface, mucilaginous, roughened, hyaline,(0.4–)1.0(–1.5) mm thick when intact, usually moreor less deteriorated in mature spores, almost alwayssloughed in older specimens (FIGS. 16–22). Layer 2permanent, semirigid, smooth, hyaline, (0.8–)1.2(–1.5) mm thick, loosely associated with layer 3

(FIGS. 16–22); in vigorously crushed spores this layerfrequently cracks and then separates from layer 3and usually protrudes because of its rigidity (FIGS. 21,22). Layer 3 laminate, smooth, hyaline to pastelyellow (3A4), (1.2–)2.0(–2.6) mm thick, sometimesstratifying into groups of or single laminae incrushed spores (FIGS. 16–22). In Melzer’s reagentonly layer 1 stains pastel red (7A5) to brownish red(10C6, FIGS. 15, 17–19). Subtending hypha hyaline topastel yellow (3A4); straight or recurved, cylindricalto slightly funnel-shaped, rarely constricted at sporebase; (4.8–)6.9(–9.8) mm wide at the spore base(FIGS. 14, 15, 18, 20–22). Wall of subtending hyphahyaline to pastel yellow (3A4); (1.8–)2.6(–3.5) mmthick at the spore base; composed of three layerscontinuous with spore wall layers 1–3 (FIGS. 21, 22).Pore (1.2–)2.5(–5.0) mm diam, open (FIGS. 21, 22).Germination unknown.

Mycorrhizal associations. In the field G. iranicumwas associated with roots of T. aestivum. In one-speciespot cultures with P. lanceolata as the host plant G.iranicum formed mycorrhizae with arbuscules andintra- and extraradical hyphae (FIGS. 23–25). Novesicles were found. Arbuscules generally were infre-quent and widely dispersed along the root fragmentsexamined. They consisted of a short trunk developedfrom a parent hypha and numerous branches with finetips (FIG. 23). Intraradical hyphae grew parallel to thelongitudinal root axis, were straight to slightly curved,and (2.3–)4.3(–7.0) mm wide (FIGS. 23–25). Theysometimes formed Y-shaped branches and coils(FIGS. 24, 25). Coils were ellipsoidal, 19.0–22.8 3

25.8–56.8 mm, when seen in a plan view, and notnumerous and not widely dispersed along rootsfragments (FIG. 25). Extraradical hyphae occurredrarely and were (2.4–)4.8(–6.8) mm wide. In 0.1%

trypan blue arbuscules stained lilac (16B5) to deepviolet (16E8), intraradical hyphae pale violet (16A3) todeep violet (16D8), coils violet (17A6–17B7), andextraradical hyphae pale violet (16A3) to grayish violet(16C6; FIGS. 23–25).

Phylogenetic position. Phylogenetic analyses of par-tial SSU sequences of nrDNA positioned G. iranicumunambiguously in Glomus group A sensu Schwarzott etal. (2001) (FIG. 13). The sequences of the speciesseparated unambiguously from described Glomusspecies. Although sequences of G. iranicum groupedtogether with a sequence obtained from roots of P.africana from Africa (Wubet et al. 2009), they arerelatively distant. The clade with G. iranicum groupedtogether without significant support with severallineages of sequences obtained from different inplanta diversity studies (Bidartondo et al. 2002; Wubetet al. 2006, 2009; Kovacs et al. 2007) but not with anydescribed Glomus species (FIG. 13).

BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1457

FIGS. 14–21. Glomus iranicum. 14, 15. Spores in loose clusters. 16–19. Spore wall layers (swl) 1–3. 20. Spore wall layers (swl)2 and 3. Note the thick swl2. Spore wall layer 1 is sloughed in this spore. 21. Spore wall layers (swl) 2 and 3 and subtendinghyphal wall layers (shwl) 2 and 3. Spore wall layer 1 and subtending hyphal wall layer 1 are sloughed in this spore. Note thecracked semirigid swl2 and the separated laminae of swl3. 14, 16. Spores in PVLG. 15, 17–21. Spores in PVLG + Melzer’sreagent. 14–21. Differential interference microscopy. Bars: 14, 15 5 20 mm; 16–21 5 10 mm.

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Specimens examined. POLAND, Szczecin, under pot-cultured P. lanceolata, 2 Jul 2009, Błaszkowski, J., 3185(HOLOTYPE, DPP); Błaszkowski, J., 3186–3207 (ISTO-TYPES, DPP) and two slides at OSC.

Etymology. Latin, iranicum, referring to Iran, theonly country in which the species has been found.

Distribution and habitat. Glomus iranicum wasfound in only one trap culture with rhizosphere soiland root fragments of T. aestivum collected from acultivated field in a weakly alkaline clay soil nearKhoramshahr, Khuzestan Province (southwesternIran; 30u309N, 48u099E) 5 Jun 1997.

Notes. The distinctive morphological characters ofG. iranicum are the hyaline to pale, small sporesformed mainly in loose clusters (FIGS. 14–17, 22) andthe permanent, semirigid, hyaline and relatively thickmiddle layer of the three-layered spore wall (FIGS. 16–22). In crushed spores this layer usually cracks,separates from the laminate layer 3 and frequentlyprotrudes because of its rigidity (FIGS. 19, 21, 22). Inaddition the fungus has a mucilaginous, short-lived

outermost spore wall layer, forming the surface, whichstains intensively in Melzer’s reagent (FIGS. 15–19, 22),and the laminate layer 3 frequently stratifies intogroups of single laminae in crushed spores (FIG. 21).

Of the described Glomus spp. forming clusters ofspores having a three-layered spore wall in which theinnermost layer is laminate, G. iranicum is mostsimilar to G. xanthium Błaszk., Blanke, Renker &Buscot (Błaszkowski et al. 2004). The darkest G.iranicum spores are pigmented similarly to thelightest G. xanthium spores. Mature spores of bothspecies have a similar size. Spore wall layer 2 in G.iranicum and G. xanthium is permanent, rigid,smooth, hyaline and of similar thickness, and thesubtending hypha of spores of both fungi has asimilar width. However most G. xanthium spores aremarkedly darker, to yellow ocher (5C7), and never arehyaline as are most spores of G. iranicum. Mostimportant at maturity spore wall layer 1 of G.xanthium generally is present as a more or lessdeteriorated, light yellow (4A4) structure (1.2–)

FIGS. 22–25. Glomus iranicum. 22. Spore wall layers (swl) 1–3 continuous with subtending hyphal wall layers (shwl) 1–3.Note highly decomposed swl1 and shwl1. 23–25. Mycorrhizae of Glomus iranicum in roots of Plantago lanceolata stained in0.1% trypan blue. 23. Arbuscule (arb) with trunk (t) developed from parent hypha (ph) and straight intraradical hypha (sih).24. Y-shaped branch. 25. Coil (c). 22–25. Spores and root fragments in PVLG. 22–25. Differential interference microscopy.Bars: 22–25 5 10 mm.

BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1459

1.8(–2.7) mm thick, which does not stain in Melzer’sreagent. In contrast spore wall layer 1 of G. iranicumis markedly thinner and stains intensively in Melzer’sreagent (FIGS. 15–19). Moreover this layer is a short-lived component of the spore wall and is rarelypresent in mature G. iranicum spores (FIGS. 17, 20–22). Finally, the subtending hypha of G. xanthiumspores is more uniform (cylindrical to flared vs.cylindrical to funnel-shaped in G. iranicum), its upperrange of width, (0.5–)0.9(–1.5) mm at spore base, doesnot attain even the lower limit of that of thesubtending hypha of G. iranicum spores, and itslumen is occluded by a septum (vs. it is open in G.iranicum; FIGS. 14, 15, 18, 20–22).

Another species of the Glomus group that sharessome characters with G. iranicum is G. viscosum T.H.Nicolson. Similarly to G. iranicum, many G. viscosumspores arise in loose clusters, the spores are hyaline topale (pale yellow; www.invam.caf.wvu.edu/, Walker etal. 1995), and their three-layered spore wall comprisestwo permanent layers (2 and 3) with nearly identicalphenotypic and biochemical properties. However G.viscosum spores generally are larger (www.invam.caf.wvu.edu/, Walker et al. 1995), and spore wall layer 2 ismuch thinner, 0.5 mm thick or less versus (0.8–)1.2(–1.5) mm thick (FIGS. 16–22) and semiflexible (www.invam.caf.wvu.edu/) versus semirigid, frequentlycracking in vigorously crushed spores (FIGS. 20–22)than G. iranicum. In addition, although spore walllayer 1 of G. viscosum is an impermanent structurelike that of G. iranicum, it does not stain in Melzer’sreagent (FIGS. 15–19), it generally remains attachedto mature spores, and it exudes a mucigel-likesubstance absorbing soil particles (www.invam.caf.wvu.edu/, Walker et al. 1995) distinguishing it fromG. iranicum.

Similar to G. africanum, G. iranicum has noapparent molecular relatives among described Glo-mus spp. (FIG. 13). Although G. iranicum shares somemorphological characters with G. viscosum and G.xanthium, the phylogenetic positions of the threespecies within Glomeromycota are different. Glomusiranicum is a member of Glomus group A, whereas G.viscosum is related to species of Glomus group B(Schwarzott et al. 2001). Although G. xanthiumbelongs to Glomus group A, its closest molecularrelatives are G. caledonium and G. mosseae (Błasz-kowski et al. 2004), which are clearly separated fromG. iranicum within the phylogenetic analyses present-ed here (FIG. 13).

ACKNOWLEDGMENTS

We thank Prof Zdzisław Koszanski, Department of WaterManagement, West Pomeranian University of Technology,

Szczecin, for collecting the A. arenaria soil-root samplefrom Varna (Bulgaria). We also thank Dr P. Schreiner,Mycologia associate editor, and two anonymous reviewersfor valuable comments. This study was supported in part bythe Polish Committee of Scientific Researches, grants 2PO4C 041 28 and 164/N-COST/2008/0, and the Hungar-ian Research Fund, OTKA K72776. A part of the study wasconducted during the Experienced Researcher Fellowshipof the Alexander von Humboldt Foundation awarded to G.M. Kovacs.

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